I. In Vitro Cell Culture
The majority of in vitro vertebrate cell cultures are grown as monolayers on an artificial substrate which is continuously bathed in a nutrient medium. The nature of the substrate on which the monolayers may be grown may be either a solid (e.g., plastic) or a semi-solid (e.g., collagen or agar). Currently, disposable plastics have become a preferred substrate for cell culture.
While the growth of cells in two-dimensions is frequently used for the preparation and examination of cultured cells in vitro, it lacks the characteristics of intact, in vivo tissue which, for example, includes cell-cell and cell-matrix interactions. Therefore, in order to characterize these functional and morphological interactions, various investigators have examined the use of three-dimensional substrates in such forms as a collagen gel (Yang et al., Cancer Res. 41:1027 (1981); Douglas et al., In Vitro 16:306 (1980); Yang et al., Proc. Nat'l Acad. Sci. 2088 (1980)), cellulose sponge (Leighton et al., J. Nat'l Cancer Inst. 12:545 (1951)), collagen-coated cellulose sponge (Leighton et al., Cancer Res. 28:286 (1968)), and GELFOAM® (Sorour et al., J. Neurosurg. 43:742 (1975)). Typically, these aforementioned three-dimensional substrates are inoculated with the cells to be cultured, which subsequently penetrate the substrate and establish a “tissue-like” histology similar to that found in vivo. Several attempts to regenerate “tissue-like” histology from dispersed monolayers of cells utilizing three-dimensional substrates have been reported. For example, three-dimensional collagen substrates have been utilized to culture a variety of cells including breast epithelium (Yang, Cancer Res. 41:1021 (1981)), vascular epithelium (Folkman et al., Nature 288:551(1980)), and hepatocytes (Sirica et al., Cancer Res. 76:3259 (1980)), however long-term culture and proliferation of cells in such systems has not yet been achieved. Prior to the present invention, a three-dimensional substrate had not been utilized in the autologous in vitro culture of cells or tissues derived from the dermis, fascia, or lamina propria.
II. Augmentation and/or Repair of Dermal and Subcutaneous Tissues
In the practice of cosmetic and reconstructive plastic surgery it is frequently necessary to employ the use of various injectable materials to augment and/or repair defects of the subcutaneous or dermal tissue, thus effecting an aesthetic result. Non-biological injectable materials (e.g., paraffin) were first utilized to correct facial contour defects as early as the late nineteenth century. However, numerous complications and the generally unsatisfactory nature of long-term aesthetic results caused the procedure to be rapidly abandoned. More recently, the use of injectable silicone became prevalent in the 1960's for the correction of minor defects, although various inherent complications also limited the use of this substance. Complications associated with the utilization of injectable liquid silicone include local and systemic inflammatory reactions, formation of scar tissue around the silicone droplets, rampant and frequently-distant unpredictable migration throughout the body, and localized tissue breakdown. Due to these potential complications, silicone is not currently approved for general clinical use. Although the original proponents of silicone injection have continued experimental programs utilizing specially manufactured Medical Grade silicone (e.g., Dow Corning MDX 4.4011®) with a limited number of subjects, it appears highly unlikely that its use will be generally adopted by the surgical community. See e.g., Spira and Rosen, Clin. Plastic Surgery 20:181 (1993); Matton et al., Aesthetic Plastic Surgery 9:133 (1985).
It has also been suggested to compound extremely small particulate species in a lubricious material and inject such combination micro-particulate media subcutaneously for both soft and hard tissue augmentation and repair, however success has been heretofore limited. For example, bioreactive materials such as hydroxyapatite or cordal granules (osteo conductive) have been utilized for the repair of hard tissue defects. Subsequent undesirable micro-particulate media migration and serious granulomatous reactions frequently occur with the injection of this material. These undesirable effects are well-documented with the use of such materials as polytetrafluoroethylene (TEFLON®) spheres of small diameter (˜90% of particles having a diameter of 30 μm) in glycerin. See e.g., Malizia et al., JAMA 251:3277 (1984). Additionally, the use of very small diameter particulate spheres (˜1-20 μm) or small elongated fibrils (˜1-30 μm in diameter) of various materials in a biocompatible fluid lubricant as injectable implant composition are disclosed in U.S. Pat. No. 4,803,075. However. while these aforementioned materials create immediate augmentation and/or repair of defects, they also have a tendency to migrate and be reabsorbed from the original injection site.
The poor results initially obtained with the use of non-biological injectable materials prompted the use of various non-immunogenic, proteinaceous materials (e.g., bovine collagen and fibrin matrices). Prior to human injection, however, the carboxyl- and amino-terminal peptides of bovine collagen must first be enzymatically-degraded, due to its highly immunogenic nature. Enzymatic degradation of bovine collagen yields a material (atelbcollagen) which can be used in limited quantities in patients pre-screened to exclude those who are immunoreactive to this substance. The methodologies involved in the preparation and clinical utilization of atelocollagen are disclosed in U.S. Pat. Nos. 3,949,073; 4,424,208; and U.S. Pat. No. 4,488,911. Atelocollagen has been marketed as ZYDERM® brand atelocollagen solution in concentrations of 35 mg/ml and 65 mg/ml. Although atelocollagen has been widely employed, the use of ZYDERM® has been associated with the development of anti-bovine antibodies in approximately 90% of patients and with overt immunologic complications in 1-3% of patients. See DeLustro et al., Plastic and Reconstructive Surgery 79:581 (1987).
Injectable atelocollagen solution also was shown to be absorbed from the injection site, without replacement by host material, within a period of weeks to months. Clinical protocols calling for repeated injections of atelocollagen are, in practice, primarily limited by the development of immunogenic reactions to the bovine collagen. In order to mitigate these limitations, bovine atelocollagen was further processed by cross-linking with 0.25% glutaraldehyde, followed by filtration and mechanical shearing through fine mesh. The methodologies involved in the preparation and clinical utilization of this material are disclosed in U.S. Pat. Nos. 4,582,640 and 4,642,117. The modified atelocollagen was marketed as ZYPLAST® brand cross-linked bovine atelocollagen. The propertied advantages of cross-linking was to provide increased resistance to host degradation, however this was off-set by an increase in solution viscosity. In addition, cross-linking of the bovine atelocollagen was found to decrease the number of host cells which infiltrated the injected collagen site. The increased viscosity, and in particular irregular increased viscosity resulting in “lumpiness,” not only rendered the material more difficult to utilize, but also made it unsuitable for use in certain circumstances. See e.g., U.S. Pat. No. 5,366,498. In addition, several investigators have reported that there is no or marginally-increased resistance to host degradation-of ZYPLAST® cross-linked bovine atelocollagen in comparison to that of the non-cross-linked ZYDERM® atelocollagen and that the overall longevity of the injected material is, at best, only 4-6 months. See e.g., Ozgentas et al., Ann. Plastic Surgery 33:171 (1994); and Matti and Nicolle, Aesthetic Plastic Surgery 14:227 (1990). Moreover, bovine atelocollagen cross-linked with glutaraldehyde may retain this agent as a high molecular weight polymer which is continuously hydrolyzed, thus facilitating the release of monomeric glutaraldehyde. The monomeric form of glutaraldehyde is detectable in body tissues for up to 6 weeks after the initial injection of the cross-linked atelocollagen. The cytotoxic effect of glutaraldehyde on in vitro fibroblast cultures is indicative of this substance not being an ideal cross-linking agent for a dermal equivalent which is eventually infiltrated by host cells and in which the bovine atelocollagen matrix is rapidly degraded, thus resulting in the release of monomeric glutaraldehyde 5 into the bodily tissues and fluids.
Similarly, chondroitin-6-sulfate (GAG), which weakly binds to collagen at neutral pH, has also been utilized to chemically modify bovine protein for tissue graft implantation. See Hansborough and Boyce, JAMA 136:2125 (1989). However, like glutaraldehyde, GAG may be released into the tissue causing unforeseen long-term effects on human subjects. GAG has been reported to increase scar tissue formation in wounds, which is to be avoided in grafts. Additionally, a reduction of collagen blood clotting capacity may also be deleterious in the application in bleeding wounds, as fibrin clot contributes to an adhesion of the graft to the surrounding tissue.
The limitations which are imposed by the immunogenicity of both modified and non-modified bovine atelocollagen have resulted in the isolation of human collagen from placenta (see e.g., U.S. Pat. No. 5,002,071) from surgical specimens (see e.g., U.S. Pat. Nos. 4,969,912 and 5,332,802); and cadaver (see e.g., U.S. Pat. No. 4,882,166). Moreover, processing of human-derived collagen by cross-linking and similar chemical modifications is also required, as human collagen is subject to analogous degradative processes as is bovine collagen. Human collagen for injection, derived from a sample of the patient's own tissue, is currently available and is marketed as AUTOLOGEN®. It should be noted, however, that there is no quantitative evidence which demonstrates that human collagen injection results in lower levels of implant degradation than that which is found with bovine collagen preparations. Furthermore, the utilization of autologous collagen preparation and injection is limited to those individuals who have previously undergone surgery, due to the fact that the initial culture from which the collagen is produced is derived is from the tissue removed during the surgical procedure. Therefore, it is evident that, although human collagen circumvents the potential for immunogenicity exhibited by bovine collagen, it fails to provide long-term therapeutic benefits and is limited to those patient who have undergone prior surgical procedures.
An additional injectable material currently in use as an alternative to atelocollagen augmentation of the subjacent dermis consists of a mixture of gelatin powder, -aminocapronic acid, and the patient's. plasma marketed as FIBREL®. See Multicenter Clinical Trial, J. Am. Acad. Dermatology 16:1155 (1987). The action of FIBREL® appears to be dependent upon the initial induction of a sclerogenic inflammatory response to the augmentation of the soft tissue via the subcutaneous injection of the material. See e.g., Gold, J. Dermatologic Surg. Oncology, 20:586 (1994). Clinical utilization of FIBREL® has been reported to often result in an overall lack of implant uniformity (i.e., “lumpiness”) and longevity, as well as complaints of patient discomfort associated with its injection. See e.g., Millikan et al., J. Dermatologic. Surg. Oncology, 17:223 (1991). Therefore, in conclusion, none of the currently utilized protein-based injectable materials appears to be totally satisfactory for the augmentation and/or repair of the subjacent dermis and soft tissue.
The various complications associated with the utilization of the aforementioned materials have prompted experimentation with the implantation (grafting) of viable, living tissue to facilitate augmentation and/or repair of the subjacent dermis and soft tissue. For example, surgical correction of various defects has been accomplished by initial removal and subsequent re-implantation of the excised adipose tissue either by injection (see e.g., Davies et al., Arch. of Otolaryngology-Head and Neck Surgery 121:95 (1995); McKinney & Pandya, Aesthetic Plastic Surgery 18:383 (1994); and Lewis, Aesthetic Plastic Surgery 17:109 (1993)) or by the larger scale surgical-implantation (see e.g., Ersck, Plastic & Reconstructive Surgery 87:219 (1991)). To perform both of the aforementioned techniques a volume of adipose tissue equal or greater than is required for the subsequent augmentation or repair procedure must be removed from the patient. Thus, for large scale repair procedures (e.g., breast reconstruction) the amount of adipose tissue which can be surgically-excised from the patient may be limiting. In addition, other frequently encountered difficulties with the aforementioned methodologies include non-uniformity of the injectate, unpredictable longevity of the aesthetic effects, and a 4-6 week period of post-injection inflammation and swelling. In contrast, in a preferred embodiment, the present invention utilizes the surgical engraftment of autologous adipocytes which have been cultured on a solid support typically derived from, but not limited to, collagen or isolated extracellular matrix. The culture may be established from a simple skin biopsy specimen and the amount of adipose tissue which can be subsequently cultured in vitro is not limited by the amount of adipose tissue initially excised from the patient.
Living skin equivalents have been examined as a methodology for the repair and/or replacement of human skin. Split thickness autographs, epidermal autographs (cultured allogenic keratinocytes), and epidermal allographs (cultured allogenic keratinocytes) have been used with a varying degree of success. However, unfortunately, these forms of treatment have all exhibited numerous disadvantages. For example, split thickness autographs generally show limited tissue expansion, require repeated surgical operations, and give rise to unfavorable aesthetic results. Epidermal autographs require long periods of time to be cultured, have a low success (“take”) rate of approximately 30-48%, frequently form spontaneous blisters, exhibit contraction to 60-70% of their original size, are vulnerable during the first 15 days of engraftment, and are of no use in situations where there is both epidermal and dermal tissue involvement. Similarly, epidermal allografts (cultured allogenic keratinocytes) exhibit many of the limitations which are inherent in the use of epidermal autographs. Additional methodologies have been examined which involve the utilization of irradiated cadaver dermis. However, this too has met with limited success due to, for example, graft rejection and unfavorable aesthetic results.
Living skin equivalents comprising a dermal layer of rodent fibroblast cells cast in soluble collagen and an epidermal layer of cultured rodent keratinocytes have been successfully grafted as allografts onto Sprague Dawley rats by Bell et al., J. Investigative Dermatology 81:2 (1983). Histological examination of the engrafted tissue revealed that the epidermal layer had fully differentiated to form desmonosomes, tonofilaments, keratohyalin, and a basement lamella. However, subsequent attempts to reproduce the living skin equivalent using human fibroblasts and keratinocytes has met with only limited success. In general, the keratinocytes failed to fully differentiate to form a basement lamella and the dermo-epidermal junction was a straight line.
The present invention includes the following methodologies for the repair and/or augmentation of various skin defects: (1) the injection of autologously cultured dermal or fascial fibroblasts into various layers of the skin or injection directly into a “pocket” created in the region to be repaired or augmented, or (2) the surgical engraftment of “strands” derived from autologous dermal and fascial fibroblasts which are cultured in such a manner as to form a three-dimensional “tissue-like” structure similar to that which is found in vivo. Moreover, the present invention also differs on a two-dimensional level in that “true” autologous culture and preparation of the cells is performed by utilization of the patient's own cells and serum for in vitro culture.
III. Vocal Cord Tissue Augmentation and/or Repair
Phonation is accomplished in humans by the passage of air past a pair of vocal cords located within the larynx. Striated muscle fibers within the larynx, comprising the constrictor muscles, function so as to vary the degree of tension in the vocal cords, thus regulating both their overall rigidity and proximity to one another to produce speech. However, when one (or both) of the vocal cords becomes totally or partially immobile, there is a diminution in the voice quality due to an inability to regulate and maintain the requisite tension and proximity of the damaged cord in relation to that of the operable cord. Vocal cord paralysis may be caused by cancer, surgical or mechanical trauma, or similar afflictions which render the vocal cord incapable of being properly tensioned by the constrictor muscles.
One therapeutic approach which has been examined to allow phonation involves the implantation or injection of biocompatible materials. It has long been recognized that a paralyzed or damaged vocal cord may be repositioned or supported so as to remain in a fixed location relative to the operable cord such that the unilateral vibration of the operable cord produces an acceptable voice pattern. Hence, various surgical methodologies have been developed which involve the formation of an opening in the thyroid cartilage and subsequently providing a means for the support and/or repositioning of the paralyzed vocal cord.
For example, injection of TEFLON® into the paralyzed vocal cord to increase its inherent “bulk” has been described. See e.g., von Leden et al., Phonosurgery 3:175 (1989). However, this procedure is now considered unacceptable due to the inability of the injected TEFLON® to close large glottic gaps, as well as its tendency to induce inflammatory reactions resulting in the formation of fibrous infiltration into the injected cord. See e.g., Maves et al., Phonosurgery: Indications and Pitfalls 98:577. (1989). Moreover, removal of the injected TEFLON® may be quite difficult should it subsequently be desired or become necessary.
Another methodology for supporting the paralyzed vocal cord which has been employed involves the utilization of a custom-fitted block of siliconized rubber (SILASTIC®). In order to ensure the proper fit of the implant, the surgeon hand carves the SILASTIC® block during the procedure in order to maximize the ability of the patient to phonate The patient is kept under local anesthesia so that he or she can produce sounds to test the positioning of the implant. Generally, the implanted blocks are formed into the shape of a wedge which is totally implanted within the thyroid cartilage or a flanged plug which can be moved back-and-forth within the opening in the thyroid cartilage to fine-tune the voice of the patient.
Although SILASTIC® implants have proved to be superior over TEFLON® injections, there are several areas of dissatisfaction with the procedure including difficulty in the carving and insertion of the block, the large amount of time required for the procedure, and a lack of an efficient methodology for locking the block in place within the thyroid cartilage. In addition, vocal cord edema, due to the prolonged nature of the procedure and repeated voice testing during the operation, may also prove problematic in obtaining optimal voice quality.
Other methodologies which have been utilized in the treatment of vocal cord paralysis and damage include GELFOAM® hydroxyapatite, and porous ceramic implants, as well as injections of silicone and collagen. See e.g., Koufman, Laryngoplastic Phonosurgery (1988). However, these materials have also proved to be less than ideal due to difficulties in the sizing and shaping of the solid implants as well as the potential for subsequent immunogenic reactions. Therefore, there still remains a need for the development of a methodology which allows the efficacious treatment of vocal cord paralysis and/or damage.